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Anc1, a Protein Associated with Multiple Transcription Complexes, Is Involved in Postreplication Repair Pathway in S. cerevisiae

Yeast strains lacking Anc1, a member of the YEATS protein family, are sensitive to several DNA damaging agents. The YEATS family includes two human genes that are common fusion partners with MLL in human acute leukemias. Anc1 is a member of seven multi-protein complexes involved in transcription, and the damage sensitivity observed in anc1Δ cells is mirrored in strains deleted for some other non-essential members of several of these complexes. Here we show that ANC1 is in the same epistasis group as SRS2 and RAD5, members of the postreplication repair (PRR) pathway, but has additive or synergistic interactions with several other members of this pathway. Although PRR is traditionally divided into an “error-prone” and an “error-free” branch, ANC1 is not epistatic with all members of either established branch, and instead defines a new error-free branch of the PRR pathway. Like several genes involved in PRR, an intact ANC1 gene significantly suppresses spontaneous mutation rates, including the expansion of (CAG)25 repeats. Anc1's role in the PRR pathway, as well as its role in suppressing triplet repeats, point to a possible mechanism for a protein of potential medical interest.

Introduction
Understanding the role of all genes that function to provide resistance upon chemical exposure will provide a systems level view of how cells respond to changing environments, and an understanding of what happens to the cell and the organism when this system is impaired. Recently, we screened all of the nonessential yeast genes to identify those that provide resistance to DNA damaging agents [1,2]. Based on the genes of known function whose deletion resulted in sensitivity, we identified several unexpected cellular processes that were overrepresented among damage sensitive mutants [1,2]. RNA polymerase II (Pol II)mediated transcription was among the many pathways that were significantly overrepresented [1,2].
The product of the ANC1 gene (also known as TAF14 and TFG3) is a member of at least seven multi-protein complexes that have distinct but related cellular roles, the common theme being their involvement in RNA Polymerase II-mediated transcription. Anc1-containing complexes include two members of the RNA Pol II holoenzyme, TFIID and TFIIF, the chromatin remodeling complexes RSC, SWI/SNF, and INO80, the histone acetyltrans-ferase complex, NuA3 and the transcriptional activation adapter complex Mediator. [3][4][5][6][7][8][9]. The sensitivity of anc1D S. cerevisiae strains to UV light, c-irradiation the DNA alkylating agents methane methylsulfonate (MMS) and 4-nitroquinoline 1-oxide (4NQO), and to hydroxyurea (HU) was recently reported [1,2,8,10]. How Anc1 promotes survival after exposure to DNA damage and replicative stress has not, until now, been explored.
The Anc1 protein contains a highly conserved YEATS domain that is present in three yeast (Yaf9, Anc1 and Sas5) and four human (ENL/MLLT1, AF9/MLLT3, GAS41 and YEATS2) proteins. Three of the four YEATS family human proteins are associated with the human mixed linkage leukemia gene: MLL gene fusions occur in ,3% of AML (acute myeloid leukemia) and 8-10% of ALL (acute lymphoid leukemia) [11]. Both ENL and AF9 are common translocation partners with MLL in these cancers, and GAS41 has been shown to interact directly with the product of the AF10 gene, another MLL fusion partner [12]. Together, ENL, AF9 and AF10 fusions with MLL account for about 35% of spontaneous human acute leukemias with MLL gene fusions [11]. The function of the YEATS domain is still largely unknown, although it was recently reported that tagged ENL binds specifically to histones H1 and H3 via its YEATS domain [13]. Moreover, the wildtype MLL protein is a member of a large multiprotein complex that contains many members of the human TFIID and SWI/SNF transcription complexes. Similar to MLL, Anc1 is a member of yeast TFIID and SWI/SNF complexes, and is thus intriguingly similar to MLL itself [14]. This, along with the fact that Anc1 and several of MLL's leukemogenic fusion partners share the YEATS domain makes Anc1 a particularly interesting candidate for mechanistic analysis.
During DNA replication, template nucleotides that have been chemically modified or lack a base altogether frequently block advancement of the replication fork, and can even cause fork collapse. Unless a stalled replication fork is enabled to restart, the cell cannot properly complete DNA replication, resulting in cell cycle arrest and cell death. The post-replication repair (PRR) pathway, exemplified by the RAD6 epistasis group in S. cerevisiae, employs a variety of mechanisms for restarting stalled replication forks. It is the least well characterized of the DNA repair pathways, and is generally divided into error-prone and error-free branches, although there is some disagreement as to the number and sub-branches therein [15][16][17][18]. The error prone branch employs specialized translesion DNA polymerases (i.e. Rev1, Pol f, Pol g) that individually, or in collaboration, allow replication past and beyond replication-blocking DNA lesions, usually in an error-prone manner. Such DNA lesion bypass enables continued replication, albeit at the cost of increased mutation, and renders the lesion available for subsequent DNA repair [19]. The error-free branch of PRR, still largely uncharacterized, competes with Rad52-mediated homologous recombination for substrates, and likely repairs these substrates by recombination between sister-strands, through either template strand switching or copy choice mechanisms [20]. The error-free branch of PRR is associated with a subset of the Rad6 epistasis group, including Rad6, Rad18, Srs2, Rad5, Ubc13 and Mms2 [18].
Rad6, an E2 ubiquitin conjugating enzyme, forms a heterodimer with Rad18, a ubiquitin ligase and single-strand DNAdependent ATPase. Under appropriate conditions the Rad6/ Rad18 heterodimer monoubiquitinates PCNA at lysine 164. PCNA thus modified activates the error-prone PRR pathway by recruiting translesion polymerases to the replication fork [21,22]. Alternately, monoubiquitinated PCNA can serve as a substrate for polyubiquitination by the Rad5/Mms2/Ubc13 complex, leading to activation of the error-free pathway instead [21,22]. Like Rad18, Rad5 is a single-strand DNA-dependent ATPase. Rad5 appears to play a complex role in these pathways, with evidence for its participation in error-prone translesion synthesis, and at least one putative branch of the error-free pathway, although its primary role is considered to be in the error-free branch [15,17,18,23,24]. It has recently been shown to have a specialized helicase activity for replication fork regression that may be important for template switching [25]. Srs2 (''Suppressor of Rad6''), a DNA-dependent ATPase and helicase, strips Rad51 from single-stranded DNA, preventing Rad51 from sequestering the DNA for homologous recombination, and allowing PRR pathway members to access the substrate, directing to toward synthesis-dependent strand annealing instead [26,27]. Thus, Srs2 acts as the gatekeeper to all of postreplicative repair, although it only suppresses damage-induced sensitivity and mutagenesis in mutants of the error-free branch of PRR [28].
In this study we investigate the basis of anc1D's sensitivity to alkylating agents. We show that ANC1 defines a new branch in the PRR pathway, one that is error-free, promotes cell survival in the presence of DNA damaging agents, and suppresses both induced and spontaneous mutation, including the expansion of CAG triplet repeats.

Analysis of Anc1-containing complexes
As discussed earlier, Anc1 is a member of several RNA Pol IIrelated multi-protein complexes, namely TFIID, TFIIF, RSC,  SWI/SNF, INO80, NuA3 and Mediator. Given these associations, we set out to determine whether the role of Anc1 in providing alkylation resistance could be assigned to one or more of these complexes, bearing in mind that Anc1 might provide resistance independently of these complexes. We therefore checked the sensitivity of mutants deleted for the non-essential members for each complex. We reasoned that if deletion mutants for other members of a particular protein complex share anc1D's damage sensitivity profile, this would pinpoint the complex via which Anc1 helps cells survive after chemical damage.
Using data from our genome-wide DNA damage sensitivity phenotyping screen, the non-essential members of Anc1's constituent complexes were checked for MMS, 4NQO and UV sensitivity [1,2] ( Figure S1). For two of the seven Anc1-containing complexes, namely TFIID and TFIIF, Anc1 is the only nonessential member, so these complexes could not be interrogated. Of the five complexes containing non-essential members in addition to Anc1, four have a majority of subunits that, when deleted, share anc1D's sensitivity to MMS, 4NQO or UV; these are Mediator, SWI/SNF, INO80, and RSC excluding only NuA3 ( Figure S1). The damage sensitivity of strains deleted for several subunits in four out of five complexes demonstrates that the role of Anc1 in survival after DNA damage is likely to be tied to the functions of at least four of its protein complexes.

Alkylation damage induces cell cycle arrest in Anc1 deficient cells
Many cell cycle-related genes are critical for survival after alkylation damage; indeed, ,45% of known cell cycle regulation genes were found to be MMS sensitive in our genome-wide screen for genes involved in damage resistance [2]. Strains mutated in genes that are necessary for the Mec1-mediated DNA damage checkpoint (i.e. MEC1, RAD53, RAD9, RAD17, RAD24) are more sensitive to killing by MMS than wildtype strains [29,30]. Given the sensitivity of the anc1D strain to MMS and 4NQO damage, it seemed plausible that their sensitivity may be due their failure to arrest in response to DNA damage [31]. To assess the effect of Anc1 on the Mec1-mediated DNA damage checkpoint, we analyzed cell cycle progression in wild-type and anc1D yeast cultures in the presence of MMS [30] (Figure 1).
As previously shown, a moderately toxic dose of MMS (0.015%) induced a Mec1-dependent S-phase arrest in wildtype S. cerevisiae [30]. The 0.015% dose of MMS used in this experiment causes minimal killing in wildtype and only moderate killing in anc1D strains (Figures 2, 3 and 4). Although anc1D strains grow more slowly than wildtype [32], the MMS-induced S-phase arrest is clearly observed in both the wildtype and anc1D strains ( Figure 1); it is important to note that no such arrest is observed in mec1D -1, rad53D, rad9D, rad17D and rad24D [29,30]. Interestingly, even when arrest is induced in several of these deletion strains, it has been found that they do not undergo normal repair [33,34]. However, anc1D cells take longer than wildtype to reach an arrested state, and also take longer to move through S phase ( Figure 1). This lag may be a result of the following: (i) anc1D's slow growth rate; (ii) a slower release from the checkpoint; or (iii) a more strongly induced cell cycle arrest (Figure 1). Comparing the untreated cell cycle profiles of anc1D and wildtype, we observed that anc1D cells spend much longer in G1 than do wildtype cells, presumably contributing to their slow growth phenotype ( Figure 1).
The reason for a prolonged G1 in anc1D cells is unclear, but may be linked to a role for Anc1 either in leaving G1 or in starting S phase. What does seem clear is that the sensitivity of anc1D cells is not due to a complete failure to arrest at the Mec1-mediated DNA damage checkpoint, although there does seem to be a delay in triggering this S-phase checkpoint [29].

Epistasis of ANC1 with established DNA Repair pathways
To determine if ANC1 functions within a canonical DNA repair pathway, we examined whether anc1D could be assigned to established DNA repair epistasis groups, namely nucleotide excision repair (here represented by rad2D), base excision repair (apn1D), homologous recombination (rad51D, rad54D), transcription coupled repair (rad26D) or postreplicative repair (PRR) (rad5D and rad6D) ( Figure 2, Figure S2). The MMS sensitivity of double mutant strains was compared to each of the single mutants as well as wildtype yeast. With the exception of rad5anc1D, the double mutants all showed additive or synergistic effects compared to the single mutants ( Figure 2, Figure S2). The sensitivity of the rad5anc1D double mutant matches that of the rad5D single mutant, indicating that ANC1 shares a genetic pathway with RAD5, a member of the postreplicative repair pathway ( Figure 2F). rad6D strains are extremely sensitive to MMS, roughly 100X more sensitive than anc1D, so we used two sets of MMS doses to observe sensitivity in this epistasis test ( Figures 3G and H). At the MMS doses to which anc1D cells begin to show sensitivity, the sensitivity of the rad6D and the rad6anc1D strains was so great that survival could not be measured. But, looking at MMS doses on a log scale, we observed an possibly epistatic relationship between ANC1 and RAD6 ( Figure 2G). A closer examination of the extremely low MMS dose range where the survival of rad6D and rad6anc1D strains can be accurately measured may, however, reveal a synergistic relationship between these two genes ( Figure 2H).
RAD5 is known to belong to the error-free branch of PRR, although studies have shown an additional role for Rad5 in the error-prone branch of the pathway [15,23,35]. RAD5 has a complex relationship with other members of the error-free branch of PRR: the rad5mms2D double mutant has an additive effects for UV or MMS induced cytotoxicity compared to the single mutants [16], and Mms2/Ubc13-dependent and -independent roles for Rad5 have recently been described [15]. RAD6, on the other hand, operates upstream of the branching between the error-prone and error-free pathways ( Figure 3A).
After establishing ANC1's genetic relationship with RAD5 and RAD6, we assayed the epistasis between ANC1 and other members of the PRR pathway, namely SRS2, MMS2, UBC13, REV3, and RAD30 ( Figure 3). The genetic relationships between the genes in the PRR pathway and the number of branches therein are a subject of some disagreement, but the pathway is generally divided into error-prone and error-free branches [15][16][17][18] (Figure 3A). Like RAD5, SRS2 was also found to be in the same genetic pathway as ANC1, with the srs2D mutation suppressing the MMS sensitivity of anc1D ( Figure 3B). The suppression of anc1D sensitivity in the srs2anc1D double mutant is consistent with earlier observations that the srs2D deletion suppresses the MMS sensitivity of several mutants in the error-free branch of the postreplicative repair pathway, including rad5D, ubc13D and mms2D [28,36]. These data support the hypothesis that Anc1 functions in the error-free branch of postreplicative repair, downstream of Srs2.
Epistasis analysis of ANC1 with RAD18 was not carried out because we were unable to create a rad18anc1D double mutant by either mating or transformation, even with Rad18 expressed from a covering plasmid during transformation. The defective alphafactor response and sporulation of anc1D have been previously noted [32]. Mutants for two other error-free pathway components, MMS2 and UBC13, showed a synergistic pattern of sensitivity to MMS when combined with the ANC1 mutation ( Figure 3C, D). From this we infer that Anc1 might act on the same type of damage as Mms2/Ubc13, but through a different pathway. The epistasis of ANC1 with RAD5 does not help us determine to which branch of PRR it belongs, as RAD5 has a role in the error-prone as well as the error-free pathway. SRS2, MMS2 and UBC13 are all . Logphase cells were diluted and plated on freshly poured MMS-containing YPD-agar plates or onto control plates with no MMS. Colonies were allowed to grow at 30uC for 2-5 days before counting. At least two replicates were counted per trial. Serial dilution and gradient plate replicates available in Figure S2. doi:10.1371/journal.pone.0003717.g002 characterized members of the error-free branch, and given the epistasis of ANC1 with SRS2, but synergistic relationship with MMS2 and UBC13, the role of ANC1 in PRR does not fit into the canonical error-free branch.
Because ANC1 is synergistic rather than epistatic with the MMS2 and UBC13 members of the error-free branch of the PRR pathway, we determined whether ANC1 lies in the error-prone pathway [37]. We analyzed the alkylation sensitivity of anc1D in combination with rev3D or rad30D, mutants in two translesion DNA polymerases involved in PRR: REV3 (with REV7) encodes DNA polymerase f, an error-prone polymerase, and RAD30 encodes polymerase g, a polymerase that is sometimes characterized as error-prone, and sometimes as error-free depending on the type of lesion being bypassed [38]. The rev3anc1D double mutant showed an additive MMS-sensitivity phenotype relative to the single mutants, indicating that Anc1 probably lies in a nonoverlapping pathway with Rev3 ( Figure 3E). The rad30anc1D double mutant, however, appeared to have synergistic sensitivity when compared to the sensitivities of the single mutants, possibly indicating a partially overlapping function between Rad30 (Pol g) and Anc1 ( Figure 3F). Thus, with respect to its genetic pathway, ANC1 appears to be independent from both the error-prone and error-free branches of postreplicative repair. Taken together, from the lack of epistasis between ANC1 and error-free branch members UBC13 and MMS2, and from the lack of epistasis with error-prone branch members REV3 and RAD30, we infer that ANC1 functions in a genetic pathway that is independent from the two established branches, and therefore, may define a new branch of PRR ( Figure 3A).
Given Anc1's presence in several transcriptionally-important complexes ( Figure S1), it seems possible that Anc1's interaction with the genes of the postreplicative repair pathway may take place on a transcriptional level. To determine whether Anc1 has an effect on the transcription of genes involved in PRR, we isolated total RNA from wildtype and anc1D cells, and assayed the transcriptional levels of the PRR genes using Affymetrix microarrays (Table 1). This analysis demonstrated that none of the PRR pathway members had significant changes in expression between the wildtype and anc1D strains, but that, as expected, the expression level of ANC1 itself was considerably and significantly lower in the anc1D strain than in wildtype (Table 1).

Induced and spontaneous mutagenesis in anc1D cells
As discussed, PRR has been divided into ''error-prone'' and ''error-free'' branches. When the error-prone pathway is impaired, cells become refractory to spontaneous and damage-induced . We made several attempts to create a rad18anc1D strain for epistasis analysis, but were unable to produce the double mutant by either mating or recombination, even in the presence of a covering plasmid bearing an intact RAD18. Log-phase cells were diluted and plated on freshly poured MMS-containing YPD-agar plates or onto control plates with no MMS. Colonies were allowed to grow at 30uC for 2-5 days before counting. Results are average of at least 2 replicates, error bars = SD, except in F.; gradient plate replica for F. in Figure S2. We were unable to create a rad18anc1D double mutant by either mating or transformation, even with Rad18 expressed from a covering plasmid during transformation. doi:10.1371/journal.pone.0003717.g003 mutagenesis; when the error-free pathway is impaired, cells become, if anything, more susceptible to damage-induced mutagenesis. Given that ANC1 is not epistatic with all members of the error-free branch of postreplicative repair, it was important to determine whether ANC1 acts in an error-free or error-prone pathway with respect to mutagenesis.
Yeast lacking Anc1 were assayed for both frameshift and point mutations as previously described [22,[39][40][41]. Disruptions of the CAN1 gene, as monitored by canavanine resistance, detected predominantly point mutations (although deletions, duplications and gross chromosomal rearrangements can also disrupt CAN1) [40] Frameshift mutations were monitored by reversion of the lys2 A 12 and A 14 alleles containing mononucleotide runs of adenines [39]. Functional LYS2 is only expressed after a 21 or +1 frameshift mutation in lys2 A 12 and lys2 A 14 , respectively [39]. Rev3 is a wellcharacterized member of the error-prone branch of PRR, and is used here as a positive control for monitoring the phenotype associated with a deficiency in an error-prone pathway (Figure 4).
ANC1 deleted cells were slightly more sensitive than wildtype yeast to UV-induced point mutations and 21 framshift mutations; in contrast, UV-induced +1 frameshift mutations were similar between anc1D and wildtype. (Figure 4A, B, C). At the CAN1 locus rad5D has been observed to result in a slight increase in UVinduced point mutagenesis compared to wildtype, although the induced mutagenesis in rad5D strains has been previously characterized as being very dependent on the mutational target being assayed [15,42,43]. This is consistent with the slight increase in induced mutagenesis observed in our anc1D strain at the CAN1 locus ( Figure 4A). In contrast, the rev3D deleted strain is refractory to UV-induced point mutations compared to both wildtype and anc1D strains ( Figure 4A). Thus, anc1D's sensitivity to damageinduced mutagenesis is consistent with Anc1 acting in an error-free rather than an error-prone pathway.
Previous studies have shown an increase in spontaneous mutation rates among mutants in the error-free branch of PRR, and a decrease in the spontaneous mutation rate among mutants in the error-prone branch [17,37,44]. Here we show that the deletion of ANC1 results in an increased frequency of spontaneous -1 frameshift mutations ( Figure 4B), and also in an increased spontaneous base pair substitution mutation rate compared to wildtype ( Figure 4D). Note that the rev3D control displays a decreased spontaneous base pair substitution mutation rate compared to wildtype. Thus, in terms of both induced and background: wildtype, anc1D and rev3D cells were exposed to UV doses as indicated. Results are mean of two replicates, +/2SD. B. and C. Induced frameshift mutagenesis in CG379-A 12 and CG379-A 14 backgrounds: WT and anc1D frameshift reversions to a functional Lys2 gene. Cells were exposed to UV doses as indicated. Results are mean of two replicates, +/2SD. D. Spontaneous point mutagenesis, +/2SD, measured as described in Materials and Methods. E. Spontaneous expansion in (CAG) 25 repeats, +/2SD, measured as described in Materials and Methods. doi:10.1371/journal.pone.0003717.g004 spontaneous mutation, the newly defined Anc1 branch of PRR is clearly error-free, protecting S. cerevisiae from both cytotoxicity and mutagenesis.

Anc1 protects against trinucleotide repeat expansions
It was recently reported that gene deletion for several members of the error-free branch of the PRR pathway, including RAD5 and SRS2, results in an expansion of CAG and CTG trinucleotide repeats (TNRs); expansion of such repeats have been associated with Huntington's disease and myotonic dystrophy [45][46][47]. In those studies, it was observed that the disease-associated TNRs, but not dinucleotide repeats or non-disease associated TNRs, are prone to expansion, but not contraction, in cells deficient in the error-free branch of the PRR pathway [45].
To determine whether Anc1 plays a role in limiting CAG expansions like other members of the error-free PRR pathway, an anc1D deletion was introduced into a strain containing a (CAG) 25 construct, and assayed for CAG expansions as described [45]. Like other PRR members, anc1D displays a statistically significant (three-fold) increase in CAG expansions compared to wildtype ( Figure 4E). This expansion is statistically indistinguishable from those in rad5D and mms2D strains, although it is considerably lower than the expansion observed for several other PRR mutants [45]. These data indicate that Anc1, like other members of the errorfree PRR pathway, plays a role in preventing the expansion of CAG trinucletide repeat sequences.

Discussion
Anc1 is known to directly interact with the catalytic protein subunits for six of the seven Anc1-containing multi-protein complexes, including TFIID, TFIIF, RSC, Ino80, SWI/SNF and NuA3 [9,48,49]. Of the six subunits with which Anc1 directly interacts, Sth1, Ino80 and Snf2 are DNA-dependent ATPases/ helicases with sequence similarity to the SNF2 family of DNAdependent ATPases [9,50,51]. The other three catalytic subunits are Tsm1 and Tgf1 that are both involved in general transcription initiation, and Sas3, the catalytic subunit of NuA3 that acetylates histone H3 [9]. Given the interaction between Anc1 and the catalytic domains of nearly all of its component complexes, plus the putative interaction between histones and the Anc1 YEATS domain, it seems likely that Anc1 acts as a regulatory adapter between chromatin and the complexes that act upon it [9,13]. The damage sensitivity of cells mutant in individual components of so many of these complexes suggests that Anc1 is involved in regulating transcription, chromatin remodeling, and as reported here, PRR, upon exposure to DNA damaging agents.
The ANC1 transcript belongs to a minority of yeast transcripts that contain a splice site. It was recently reported that ANC1 mRNA splicing is regulated by Cdc40, a protein involved in controlling cell cycle progression [31]. In the absence of CDC40, cells arrest in G2/M, and the addition of intronless ANC1 cDNA only partially mitigates this arrest [31,52] indicating that Cdc40 may have other splicing targets in addition to the ANC1 mRNA, or may have yet another function. The slow transition out of G1 that we observed in anc1D cells is also observed in cdc40D cells [52,53], and like anc1D, cdc40D mutants are sensitive to a variety of DNA damaging agents, including hydroxyurea, MMS, 4NQO and UV [2,53]. However, the sensitivity of cdc40D cells to MMS or HU is not suppressed when intron-less ANC1 cDNA is expressed [53]. Of relevance to this study, a temperature sensitive allele of cdc40D was shown to be epistatic to an allele of rad6D in terms of MMS sensitivity during log phase growth, although neither allele was characterized as being null [54]. Since a correctly spliced ANC1 transcript does not suppress the MMS or HU sensitivity of cdc40D cells, we must conclude that Cdc40 has another function in allowing cells to survive after DNA damage that is independent from ANC1 transcript splicing. Like ANC1, UBC13 and MMS2 are intron-containing genes in the PRR pathway [55]. Given the observed epistasis between alleles of cdc40D and rad6D after MMS treatment [54], and the failure of the correctly spliced ANC1 transcript to complement a cdc40D mutant's damage sensitivity [53], it is worth exploring whether Cdc40 mediates the splicing of the MMS2 and/or UBC13 transcripts as well.
Several pieces of evidence support a role for Anc1 in the PRR pathway. Based on the suppression of anc1D's sensitivity by srs2D, ANC1 can be placed genetically downstream of SRS2, as was previously observed for other members of the error-free PRR pathway [28,35,36]. ANC1 also shares a genetic pathway with RAD5, a downstream member of the error-free pathway and possibly with RAD6, which lies between SRS2 and RAD5 in the genetic model of the PRR pathway ( Figure 3A). The synergism observed at low MMS doses between rad6D and anc1D may imply a role for Anc1 that is partially parallel to that of Rad6, possibly indicating that Anc1 is involved in one of the several functions of Rad6. The lack of epistasis between ANC1 and other error-free branch members MMS2 and UBC13 provides evidence for a new, Mms2/Ubc13 independent branch of the PRR pathway. Given that we were unable to create a rad18anc1D double mutant by mating or transformation, even in the presence of a covering plasmid bearing an intact RAD18, we do not yet know whether Rad18 also plays a role in the new pathway defined by Anc1; however, since no Rad6-independent role for Rad18 has been described, it seems likely that Rad18 also plays a role in the Anc1branch of PRR.
Two types of mutagenesis data indicated that the Anc1containing branch of the PRR pathway deals with DNA damage in an error-free manner. First, the ANC1 deletion, similar to deletions for most members of the error-free PRR pathway [28,43], causes an increase in both induced and spontaneous point mutation compared to wildtype. Second, anc1D mutants display a significant increase in the expansion of CAG tri-nucleotide repeats compared with wildtype, a trait that was recently identified in all of the tested members of the error-free branch of the PRR pathway, including srs2D and rad5D [45]. These mutagenesis data are consistent with a role for Anc1 in error-free PRR.
The role of Anc1 in PRR may be crucial for understanding the interaction of key players in the cellular response to DNA damage. Anc1 interacts physically with Mus81, a structure-specific endonuclease in the XPF family involved in cleaving stalled replication forks [56,57]. Mus81 forms a heterodimer with Mms4 for its endonuclease activity, and deletions of either partner results in sensitivity to MMS and 4NQO [1,2]. Mus81 is speculated to be involved with the PRR pathway (in addition to its better characterized role in homologous recombination) by means of its cleavage of the stalled replication forks that the PRR pathway acts upon [57]. Furthermore, there is genetic evidence in S. pombe that srs2D and mus81D are epistatic with respect to their MMS, UV and HU sensitivities [58], although in S. cerevisiae the mms4srs2D double mutant displays a synergistic effect compared with either of the single mutants after MMS or UV treatment, suggesting that their pathways may partially overlap [59] Having demonstrated the membership of ANC1 in the error-free branch of PRR, it seems likely that the physical interaction between Mus81 and Anc1 relates to Mus81's cleavage function in PRR. The method by which Mus81 recognizes its substrates is not well understood, but it seems possible that Anc1, through its presumed interaction with histones [13], allows the Mus81 endonuclease access to sites where its cleavage will initiate the sister-strand recombination that drives error-free PRR.
Given the direct interaction between the YEATS domain of ENL with histones H1 and H3, and the interaction of Anc1 with the catalytic subunits of so many transcriptionally-important complexes [9,13], it may be hypothesized that Anc1 acts as a DNA-damage mediated adapter between chromatin, transcription and PRR repair at or near sites of DNA damage. Since transcription generally continues through S-phase, while DNA is being replicated, the collision of the transcriptional machinery and stalled replication forks is thought to be a common event [60]. In recent years there has been considerable interest in the phenomena of transcription-associated mutation (TAM) and transcription-associated recombination (TAR), which characterize the mutagenesis and recombination that occur when the transcription and replication machineries collide [60]. Mediation of the interaction between these machineries by a common member (Anc1) of the transcription complexes is a possibility worthy of further exploration. It is possible that the new branch of postreplicative repair represented by Anc1 is responsible for mediating the repair of replication forks that have stalled as a result of the collision between transcription and replication machineries. Furthermore, the role of the human YEATS containing leukemia-associated proteins, ENL, AF9 and GAS41, in both the human post-replication repair pathway, and in polyglutamine expansions such as those associated with Huntington's disease is certainly worthy of further exploration, and may provide insight into the molecular basis of such disparate diseases as leukemia and Huntington's disease.

Yeast Strains and Media
Yeast strains used in this study are listed in Table 2. Yeast strains were grown in standard media, including YPD and synthetic complete (SC) medium. All strains are congenic with the BY4741 background (MATa his3D1 leu2D0 met15D0 ura3D0), except for the spontaneous mutagenesis and the trinucleotide repeat assays as specified below in Induced and Spontaneous Mutagenesis Assays and Table 2. Double mutants were created by transformation of an anc1D::URA3 linear cassette into G418 resistant strains from the genome-wide deletion collection (Invitrogen-ResGen) [61,62]. Homologous ends allowed the cassette to recombine into the endogenous location of ANC1 [61]. Constructs were confirmed by PCR and/or DNA sequencing.

Flow Cytometry
Log phase cells were washed twice in 50 mM Tris pH 7.8, resuspended in 50mM Tris pH 7.8 containing RNase A (1 mg/ ml) and incubated at 37uC overnight. Cells were pelleted and resuspended in 55 mM HCl containing 5 mg/ml Proteinase K, incubated at 37uC for 30 min, washed once with 200mM Tris pH 7.5, 211 mM NaCl, 78mM MgCl 2 , then resuspended in the same buffer with 1mg/ml of propidium iodide before assaying by flow cytometry using a FACScan cytometer (Becton Dickinson) and CellQuest Pro software. Two independent assays were performed to confirm reproducibility, and analysis was performed using FlowJo software Version 6.4.7.
Sensitivity of deletion strains to DNA damaging agents in the genome-wide screen The sensitivity of every non-essential gene deletant in S. cerevisiae was previously determined [1,2]. Relative sensitivity values were generated using a scoring scheme that allocated values of 4, 3, 2, or 1 depending on the concentration of agent where strain sensitivity was identified; 4 is allocated to the lowest, and 1 is allocated to the highest concentration of damaging agent in the plate. These values were allowed to accumulate in each replicate, and then they were summed across all replicates. For example, in replicate 1, strains sensitive to all concentrations of agents received a score of 10 (4+3+2+1), and this was summed over all 3 replicates for a final score of 30 (10+10+10). Damage-sensitive strains had scores that ranged from 30 (most sensitive) to 2 (least sensitive) [2]. All data is available at http://genomicphenotyping.mit.edu/ newpages/source2.html.

RNA Extraction
Three independent colonies of both wildtype and anc1::G418 R were grown overnight, then diluted and grown into log phase for 4-5 hours in YPD. RNA was extracted using Qiagen's RNeasy Mini Kit, checked for quality using an AgilentBioanalyzer and 20 ug of total RNA were hybridized using Affymetrix eukaryotic labeling protocols on Affymetrix YG-S98 microarrays (Cogenics, North Carolina).

Analysis of Microarray Data
Repair proficient and deficient strains were analyzed in triplicate on YG-S98 arrays. Normalization was carried out using the Robust Multichip Average (RMA) algorithm [63]. Arrays were analyzed using Microarray Suite 5.0 to obtain Absent/Present calls and filtered for transcripts that were not expressed in any experiment. Differential gene expression was calculated using a dual filtering criteria; (1) an estimation of statistical significance through the Local Pooled Error test (LPE) [64] calculated using S-Plus Array Analyzer with an adjustment for false discovery rate calculation of p value of ,0.05 (Benjamini Hochberg) and (2) a fold change (FC) limit of 1.5. The raw data files from this experiment have been submitted to the Gene Expression Omnibus Database (www.ncbi.nlm.nih.gov/geo), accession number GSE12150.

Survival Curves/Epistasis Assays
Log-phase cells were diluted and plated on freshly poured MMS-containing YPD-agar plates or onto control plates with no MMS. Colonies were allowed to grow at 30uC for 2-5 days, depending on rate of growth for each strain, and survival was calculated by dividing the number of surviving colonies at a given MMS dose by the number of colonies that grew in the untreated sample. At least two replicates were counted per trial.

Induced and Spontaneous Mutagenesis Assays
Yeast strains CG379-A 12 and CG379-A 14 from [39] revert by 21 and +1 frameshifts in LYS2::InsE-A 12 and LYS2::InsE-A 14 , respectively, were used to measure frameshift mutation frequencies. These strains are isogenic with CG379 (MATa ade5D1 his7D2 leu2D3, 112 trpD289 uraD52) [39]. Frameshifts were calculated by comparing the number of Lys + revertants growing on Lys 2 media to the number of colonies on a YPD control. Point mutation frequencies were measured in a BY4741 background. Canavanine-resistant mutations were measured on synthetic complete medium containing 0.004% (or 30 mg/liter) canavanine [40]. In the induced mutagenesis assay, UV doses of 0, 7, 14 and 21 J/m 2 were administered using a UV Stratalinker 2400 (Stratagene). Cells were grown into log phase, then serially diluted and plated onto YPD or Canavanine containing plates before exposure to UV. Colony formation on YPD was used to calculate the total number of cells plated on canavanine-containing plates, for a final calculation of mutants per 10 7 survivors.
In the spontaneous mutagenesis assay, forward mutations at CAN1 were determined based on the protocol previously described in Glassner et al. [41]. Briefly, an overnight culture of each strain was diluted to 4000 cells/ml in 5mL of YPD in 10 cultures. The cultures were allowed to grow at 30uC for 5 days, then a small amount diluted 10 52 fold on YPD to assay for viable cells, and the remainder concentrated to 1 mL, and 100ul plated on 0.04% Canavanine-containing synthetic complete medium to assay for Can R mutants. Mutagenesis rates were calculated using the Drake Formula [65]. Figure S1 Sensitivity of strains deleted for non-essential members of Anc1-containing complexes to MMS and 4NQO and UV. Values for increasing sensitivities from 2-30 were calculated as described in Materials and Methods, by Begley et al, 2004 and as displayed at http://genomicphenotyping.mit.edu/ newpages/source2.html. A) Mediator complex, B) SWI/SNF complex, C) Ino80 complex, D) RSC complex-although RSC14 is not essential, there is no sensitivity data available, E) NuA3 complex.


extfieldsofstudy: Biology Medicine

text:
Gene rearrangement analysis and ancestral order inference from chloroplast genomes with inverted repeat

Background Genome evolution is shaped not only by nucleotide substitutions, but also by structural changes including gene and genome duplications, insertions, deletions and gene order rearrangements. The most popular methods for reconstructing phylogeny from genome rearrangements include GRAPPA and MGR. However these methods are limited to cases where equal gene content or few deletions can be assumed. Since conserved duplicated regions are present in many chloroplast genomes, the inference of inverted repeats is needed in chloroplast phylogeny analysis and ancestral genome reconstruction. Results We extend GRAPPA and develop a new method GRAPPA-IR to handle chloroplast genomes. A test of GRAPPA-IR using divergent chloroplast genomes from land plants and green algae recovers the phylogeny congruent with prior studies, while analysis that do not consider IR structure fail to obtain the accepted topology. Our extensive simulation study also confirms that GRAPPA has better accuracy then the existing methods. Conclusions Tests on a biological and simulated dataset show GRAPPA-IR can accurately recover the genome phylogeny as well as ancestral gene orders. Close analysis of the ancestral genome structure suggests that genome rearrangement in chloroplasts is probably limited by inverted repeats with a conserved core region. In addition, the boundaries of inverted repeats are hot spots for gene duplications or deletions. The new GRAPPA-IR is available from .

Background
Mutations in a genome consist of not only base pair level changes but also events that alter the chromosome structure, such as inversions, duplications or deletions [1]. Ancestral gene sequence inference has led to significant predictions of protein functional shift and positive selection [2]. For example, comparisons of orthologous chromosomal segments showed heterogeneous rates of evolution of the X chromosome in human, mouse and rat [3]. However, on the genome level, the evolutionary change of genome structure is poorly understood. Inference of ancestral genomes was mainly achieved at the DNA level, but limited to closely related organisms where rearrangements were negligible, partly because of the complexity in assigning genes in duplicated segments to orthologous groups [4]. In this paper, we take a simple, alternative data set of chloroplast genomes to study the phylogeny and genome structural changes. Chloroplasts are the green, photosynthetic organelles that originated from a free-living cyanobacteria-like ancestor [5]. Chloroplasts maintained a reduced genome through over one billion years of endosymbiosis [6]. Typical chloroplast genomes are circular single chromosomes with 120 -200 genes, which encode proteins, tRNAs, rRNAs and hypothetical open reading frames. Most chloroplast genomes consist of four distinct parts: two duplicated regions (inverted repeats, or IR) separated by a large single copy (SSC) and a small single copy (ISC) region. Figure 1 shows the four regions (LSC-IRa-SSC-IRb) of Gossypium hirsutum chloroplast [7]. One common characteristic of the chloroplast IR is the presence of three rRNA genes (rrn5s, rrn16s and rrn23s, or rrf, rrs, and rrl), which are homologous to genes of the cyanobacterial rrn operon. The structure of chloroplast genomes of land plants is highly conserved, with almost collinear gene order, except for elevated level of rearrangements in specific lineages including green algae [8], conifers and members of a few flowering plant families including Campanulaceae [9], Geraniaceae [10] and Fabaceae [11]. The gene content of chloroplast IRs vary greatly, largely due to the expansion and contraction of the IR at the IR-SC boundaries; this "ebb and flow" of IR boundary has been observed even within a genus [12,13]. Chloroplast genomes of green algae (charophyte and chlorophyte algae) also contain more variations of gene order and some are highly rearranged [8], with some evidence that rearrangements may be adaptive in nature [37]. Because of their compact size and the availability of conserved DNA probes, many chloroplast genomes have been mapped [14], and 121 have been completely sequenced to date. Thus, chloroplast genomes provide an ideal example for modeling genome rearrangements over a broad evolutionary time scale.

Genome rearrangements
We assume a reference set of n genes {g 1 , g 2 , • • •, g n }, thus a genome can be represented as a signed ordering of these genes, and each gene is given an orientation that is either positive, written g i , or negative, written -g i . Genomes can evolve through events such as inversions, transpositions and transversion, as well as other events such as deletions, insertions and duplications of fragments. Let G be the genome with signed ordering of g1, g2,…, g n . An inversion (reversal) between indices i and j (i ≤ j), transforms G to a new genome with linear ordering A transposition on genome G acts on three indices i,j,k, with i ≤ j and k &owm; [i,j], picking up the interval gi,gi+1, … ,gj and inserting it immediately after gk. Thus genome G is replaced by (assume k > j): Because gene rearrangements are rare genomic events [15], gene-order data enable the reconstruction of evolutionary events far back in time, thus many biologists have embraced this new source of data in their phylogenetic work [9,14,16,38].

Genome rearrangement analysis
Given two genomes G 1 and G 2 , we define the edit distance d(G 1 , G 2 ) as the minimum number of events required to transform one genome into the other. The breakpoint distance [17] is not a direct evolutionary distance measurement. A breakpoint in G 1 is defined as an ordered pair of genes (g i , g j ) such that g i and g j are adjacent in G 1 but not in G 2 . The breakpoint distance is simply the number of breakpoints in G 1 relative to G 2 . When only inversions are allowed, the edit distance is the inversion distance. Hannenhalli and Pevzner [18] developed a mathematical and computational framework for signed gene-orders and provided a polynomial-time algorithm to compute the edit distance between two signed gene-orders under inversions; Bader et al. [19] later showed that this edit distance can be computed in linear time. However, computing the inversion distance is NP-hard in the unsigned case [20].
Gene rearrangement phylogeny was first proposed by Sankoff and an algorithm using break-point distance was implemented in BPAnalysis [4]. The inversion phylogeny was introduced to improve the accuracy and was implemented in GRAPPA. Extensive simulations showed that inversion median were superior to breakpoint median [21] and the trees returned were more accurate than using either distance-based or parsimony methods [22]. The current version of GRAPPA (version 2.0) is able to estimate the phylogeny and inversion medians using − + + + genomes with equal gene content (i.e., no insertion, deletion or duplication are allowed) [22]. A scaled-up version, DCM-GRAPPA, is able to estimate the gene-order phylogeny with very high accuracy for thousands of genomes, thus greatly increasing the power of genome phylogeny using large datasets [23]. We extended GRAPPA [24] so that it is able to analyze data sets with a limited number of deletions, but no duplication is allowed. To remedy this problem, we develop a new algorithm (GRAPPA-IR) for chloroplast genomes that take into account the unique quadripartite structure (e.g., LSC-IRa-SSC-IRb), which is common to not only the chloroplast genomes, but also some other IR-containing DNAs. The assumption of our new approach is that inversions do not occur across inverted repeats, because the genome structure will be disrupted by such inversions that "flip" the repeats from inverted to the same orientation. According to the model, a change of gene content within the IR region is mainly due to growth or shrinkage of the IR at the IR-SC boundaries. This approach is in agreement with the observation Gene map of the Gossypium hirsutum chloroplast genome Figure 1 Gene map of the Gossypium hirsutum chloroplast genome. The bright thick lines indicate the extent of the inverted repeats (IRa and IRb), which separate the genome into small (SSC) and large (LSC) single copy regions. The gene map is drawn using CGView [35].
that in most IR-containing chloroplasts, the gene content in the whole genome is conserved, but IR length and IR gene contents can be varied.

Results and discussion
We assess the new GRAPPA-IR through both biological and simulated datasets.
A reference phylogenetic tree was constructed using the maximum parsimony method with 50 concatenated proteins. Cyanophora proteins were used to root the tree. The reference phylogeny of these six chloroplast genomes is shown in Figure 2. The reference tree is the same as the phylogeny by Lemieux et al. [25] in which Mesostigma is basal to other green plants and algae. An alternative phylogeny was published by Karol et al. based on maximum likelihood analysis of four chloroplast genes and including many more algal taxa, in which Mesostigma is basal to charophyte green algae and sister to chlorophyte green algae [26]. We extracted 73 unique genes from the six genomes. Actual number of genes included in each genome ranges from 76 to 80 due to duplicated genes in AIR. The gene set includes 62 characterized protein-coding genes, 3 rRNAs, 7 tRNAs (identified by amino acid anticodons) and a hypothetical conserved open reading frame (ycf1). The encoding reflects the order and orientation of genes in the genome. Location of multi-exon genes was determined by the starting position of the first exon.
In one case, the order of overlapping genes (psbD-psbC) was determined by the position of the start codon. We evaluate all possible trees for the six genomes using the GRAPPA-IR. After 100 min of computation on a PIV 3.4GHz workstation, GRAPPA-AIR returned a best tree with 74 inversions, with a topology agrees with the reference tree, which is shown in Figure 3.
We tested this data set with the original GRAPPA by ignoring the region boundaries and removing the IRb region. The inference allows inversions to occur across the IR and single copy regions. The best tree obtained requires 73 inversions, yet the topology ( Figure 4) is very different from the reference trees and is in conflict with the biological relationship of these organisms. Although GRAPPA is a heuristic, extensive testing on simulated and biological data confirmed its high accuracy, thus its failure in this test suggests that the IR-boundary do perform a unique role in the evolution of chloroplast genomes, and a better method as GRAPPA-IR should be preferred.
We examined the reconstructed ancestral gene orders on the best tree returned by GRAPPA-IR. This six-genome dataset contains chloroplast genomes of land plants, green algae and a flagellate protist, which were separated by at least 450 million years of evolution [27]. The ancestral chloroplast genomes of land plants and algae contain inverted repeats, which is consistent with the hypothesis that IR is a feature derived early in the chloroplast endosymbiosis [16]. Although the sequenced cyanobacteria Nostoc and Synechococcus do not maintain rrn-containing IRs, if other cyanobacteria are identified with structures similar to the chloroplast IR, then it would suggest an even earlier origin for this structure. In addition, the ancestral IR contains the same gene content to that of Mesostigma, which agrees with the observation that Mesostigma chloroplast genome encodes several ancestral gene clusters [25]. By comparison of ancestral gene orders to the extant genomes, it is possible to test formally the evolutionary force of gene order changes. For example, The reference phylogeny of chloroplast genomes from land plants and green algae Figure 2 The reference phylogeny of chloroplast genomes from land plants and green algae.

Nephroselmis (no)
Chaetosphaeridium (cg) The best tree obtained by GRAPPA-IR. The topology is the same as the reference tree Figure 3 The best tree obtained by GRAPPA-IR. The topology is the same as the reference tree.

Psilotum (pn)
ancestral gene clusters may be more likely to be maintained if they share related function and are under constraints in the face of genome rearrangements.

Simulations to assess accuracy
Phylogenetic analysis methods deal with lost historic information, thus their accuracy should also be assessed through simulations, where the true evolutionary history is known.
We first define our measure for the accuracy of reconstructed trees. Given an inferred tree, we compare its "topological accuracy" by computing "false negatives" and "false positives" with respect to the "true tree" [28]. For every tree there is a natural association between every edge and the bipartition on the leaf set induced by deleting the edge from the tree.
Let T be the true tree and let T' be the inferred tree. An edge e in T is "missing" in T' if T' does not contain an edge defining the same bipartition; such an edge is called a false negative (FN). Note that the external edges (i.e. edges incident to a leaf) are trivial in the sense that they are present in every tree with the same set of leaves. The false negative rate is the number of false negative edges in T' with respect to T divided by the number of internal edges in T. The false positive (FP) rate is defined similarly, by swapping T and T'. The Robinson-Foulds (RF) rate is defined as the average of the FN and FP rates.
In this study, we generated datasets of 6 and 10 genomes, each with 78 genes (70 genes in LSC, 5 in SSC and 3 in IR), roughly in the range of our dataset described in the paper. We used a large range of evolutionary rates: let r denote the expected number of evolutionary events along an edge of the model tree, we used values of r in the range of 4-10. The actual number of inversions along each edge is sampled from a uniform distribution on the set {1,2,…, 2r}. Given the model tree, we assigned the identity gene order to the root, and randomly generated gene order for each node based on the edge length and the gene order of its parent, with the assumption that inversions can not cross the IR boundaries. For each combination of parameter settings, we simulated 20 datasets and averaged the results.
We compared GRAPPA-IR to the original GRAPPA. We considered all trees with the minimum score given by both methods and took their strict consensus (of course, most time there is only one single best tree was returned). Therefore, the trees returned by both methods need not to be fully resolved and they tend to have somewhat better rates for false positives than for false negatives. As a result, we only report FN rates rather than FP rates or a single Robinson-Foulds score [29]. Figure 5 and Figure 6 show the results. This simulation indicates that GRAPPA-IR is clearly more accurate than the original GRAPPA for datasets with r < 10, which is in accordance with results on the six-genome dataset.
Since the inferred ancestral genomes have direct impact on the tree scores, we can examine the quality of the inferred ancestral genomes by comparing the best tree score returned by GRAPPA-IR with the known true tree score. Averagely speaking, scores of the best trees returned by GRAPPA-IR was only about 7% less than the true scores, which strongly indicates that the inferred ancestral gene orders should be very close to the true ancestral genomes.
FN rate for GRAPPA-IR (solid line) and GRAPPA (dashed line) as a function of the evolutionary rate r for 6 genomes  The best tree when no IR boundary is imposed, with score 73 Figure 4 The best tree when no IR boundary is imposed, with score 73. Notice that the topology is different from the reference tree.

Mechanisms of IR expansion
The gene content of the IR varies across land plants, even in a single genus or family [12]. It is known that homologous recombination is frequent between the two copies of IR [30]. In a single chloroplast, hundreds of copies of chloroplast DNA co-exist as circular monomer, dimer and linear chromosomes. In the cellular endosymbiosis environment, the selection on accuracy of replication may have been relaxed to the degree that unequal recombination and replication slippage contribute to the expansion or shrinkage of IRs. Short repeat motifs may facilitate inter-molecular recombination and create diversity of chloroplast genomes in a population [31]. On the other hand, the intra-molecular recombination process should homogenize the sequences of the two IRs and thus the particular IR size and the gene content are maintained. The two counteracting phenomena may have played important roles in shaping the current diversity of chloroplast genome gene orders.

Duplications and genome stability
We found that incorrect gene order phylogenies were recovered without considering the IR boundary information. This suggests that maintenance of the IR is necessary in the evolution of chloroplast genomes in most of the cases. We propose that IR provides an insulation mechanism that stabilizes the genome structure, and the genes in single copy regions do not commute across the IR. This agrees with the observation that gene rearrangements are more frequent in chloroplast genomes without IR [32]. However, some genomes with residual IRs but infrequent gene movements between single copy regions compared to related lineages do not conform to the hypothesis [9]. Future experimental studies on highly rearranged chloroplast genomes, for example, in the green alga Chlamydomonas lineage [8,37], may shed light on the maintenance of IR and genome rearrangements.

Comparison to other methods
A similar approach used for human and mouse genome comparison showed the optimal sorting of X chromosomes by at least 7 inversions [33]. This is a moderate amount of change compared to the level we observe in many chloroplast genomes. If duplications and deletions are considered in a finer scale, the process will be much complex, as suggested by the reconstruction of one 1.1 Mb region in the eutherian mammal ancestor [4]. All methods proposed in this paper could be applied to MGR and other method, which will result in a whole new set of tools for botanists interested in genome level evolution.

Conclusions
We implement a new method to infer phylogeny and ancestral gene orders with inverted repeats. Tests on a biological and simulated dataset show GRAPPA-IR can accurately recover the genome phylogeny as well as ancestral gene orders. Close analysis of the ancestral genome structure suggests that genome rearrangement in chloroplasts is probably limited by inverted repeats with a conserved core region. In addition, the boundaries of inverted repeats are hot spots for gene duplications or deletions. This analysis provides new insight into the genome evolutionary process.

Methods
We developed a new method (GRAPPA-IR) to handle inverted repeats. This method is available online from http://phylo.cse.sc.edu.

Mapping contents in ancestral genomes
The first step of our new method-GRAPPA-IR is to determine the gene contents for each region in the genomes involved. Based on our previous research [24], such mapping can dramatically reduce the search space and improve the overall accuracy. When a genome is on a leaf (i.e., it is an extant taxon), we can easily determine the gene content for the LSC, ISC and IR regions through direct observation. However, since we do not know the gene order at each internal genome, we can only estimate the gene content for each region based on the assumption that all evolutionary events that alter the gene order are rare and that concurrent (i.e., parallel) changes in two children are less likely than a change in the parent. Thus, at each internal node, for a given region, when the regional gene contents for the two children are known, we face three possibilities of assigning a gene to the region:  FN rate (%) evolutionary rate r 1. If both children have gene g in the same region, then the parent has g in that region; otherwise, both children need to expand (or shrink) IRs and include g in that region, with a very low probability.
2. If neither child has g, then g is absent in the parent. Since the genomes we test all share 70 unique genes, we do not consider this case.
3. If g is located in different regions between the children, then it could be in either region of the parent. The two choices are equally likely without further information from the phylogeny. If the tree is rooted, we use the gene content in the evolutionary path to break the tie; otherwise, we are left with an undetermined outcome for g.
If a gene is undetermined in some internal node, it may become determined through a propagation of decisions from the leaves to the root. The estimated gene content for the internal nodes of the reference tree is presented in Figure 7. This figure shows that the gene contents of the IR and ISC regions vary among the genomes. However the gene order of part of IR is highly conserved. For example, some genes (rrn5, rrn16, rrn23, coded as 67, 68 and 69, respectively) are always kept together in the IR. If we assume that inversions do not cross the IR boundary in most chloroplast genomes, then the evolution of chloroplast genome structure can be hypothesize as undergoing the following two steps: 1. A genome was divided into regions and inversions occurred independently in each region.

2.
A segment from single copy regions was copied twice and joined to the existing inverted repeats, and the new genomes with longer IRs propagated.
One should notice that the above two steps could happen several times along each edge. Based on the above assumption, we can further simplify the gene content of IR and SSC, so that in the evolutionary path, IR regions for all genomes (leaves and internal) contain gene (67 68 69), and the SSC regions contain gene (70 71 72 73). This operation treats duplicate genes at the boundaries of IR and SSC as the last step towards the observed gene orders in the evolutionary path. Thus it is possible to ignore the duplications and reduce the problem to all leaf genomes of equal gene content (or with deletions). The simplified gene content map is shown in Figure 8.

Phylogeny and ancestral genome reconstruction
We can reconstruct the phylogeny after the regional gene contents of all genomes are determined. Since the gene contents are reduced to equal after the simplification step, it is feasible to use GRAPPA to infer an inversion phylogeny. That's to say, if inversions are allowed to cross the boundaries of pre-determined IR and single copy regions, we can use the original GRAPPA to compute the ancestral gene orders and the phylogeny. However, this is unlikely, since we do not observe any inversions involving genes in both the IR and single copy regions. Based on our proposed model, we assume that all inversions are bounded by the boundary of IRs.
Bounding inversions into each region makes each region (LSC, SSC and IR) independent, which greatly simplify the computation. For two genomes G 1 and G 2 , the genomic distance between these two can be defined as d(G 1 ,G 2 ) = d(SSC 1 , SSC 2 ) + d(LSC 1 , LSC 2 ) + d(IR 1 , IR 2 ), i.e., the overall distance is the summation of all regional distances. If no gene content is changed for each region, then the regional distance can be computed using the linear algorithm proposed by [19]. However, if the regional content is not equal (deletions or insertions occur), then more complex algorithm [34] should be used. The small-  est binary tree contains only three leaf genomes and one ancestor. This special case is called median problem in some literatures. Given three genomes G 1 , G 2 and G 3 , solving a median problem is to find a genome G 0 that can minimize the sum of distances from itself to the three given genomes. Although it is the simplest case for multiple genome analysis, it is proved to be NP hard even for the simplest distance of breakpoint. Since we can deal with each region independently, the median problem can also be divided into three regional median problems, each of which is constructed from genes in the same region of genomes g 1 , g 2 and g 3 . After the regional median is obtained, the median solution on the whole genome can be constructed simply by concatenating the regions together. Again, the regional median problems can be solved using any of the available inversion median solvers, such as [20] (for equal regional gene content) or [24] (with deletions and insertions).
To analyze a dataset with more than three genomes, GRAPPA-IR uses an exhaustive approach devised for the original GRAPPA-it must test all possible trees to find the one with the minimum number of inversions. For each tree, the program tests a lower bound [22] to determine whether the tree is worth scoring; if so, then the program will iteratively solve the median problems at internal nodes and update the internal genomes, until no change occurs, as outlined in Figure 9. The tree with the lowest score (smallest number of changes) will be returned as the phylogeny, and internal gene orders with respect to the phylogeny can be treated as the estimated ancestral genomes.
The scoring procedure of GRAPPA-IR when the tree is given Figure 9 The scoring procedure of GRAPPA-IR when the tree is given.


s2fieldsofstudy:
Biology
Environmental Science

text:
Colouration and Colour Changes of the Fiddler Crab, Uca capricornis: A Descriptive Study

Colour changes in animals may be triggered by a variety of social and environmental factors and may occur over a matter of seconds or months. Crustaceans, like fiddler crabs (genus Uca), are particularly adept at changing their colour and have been the focus of numerous studies. However, few of these studies have attempted to quantitatively describe the individual variation in colour and pattern or their adaptive significance. This paper quantitatively describes the colour patterns of the fiddler crab Uca capricornis and their ability to change on a socially significant timescale. The most dramatic changes in colour pattern are associated with moulting. These ontogenetic changes result in a general reduction of the colour pattern with increasing size, although females are more colourful and variable than similarly-sized males. Uca capricornis are also capable of rapid colour changes in response to stress, but show no endogenous rhythms associated with the semilunar and tidal cycles commonly reported in other fiddler crabs. The extreme colour polymorphism and the relative stability of the colour patterns in Uca capricornis are consistent with their use in visually mediated mate recognition.

Introduction
Many animals are capable of altering their colouration. These changes may take place over a matter of minutes or seconds, and are often involved in social interactions [1,2], background matching [3], or thermoregulation [4]. Colour changes can also occur on a longer time scale; in response to seasonal changes in the environment [5] or in association with displays during the breeding season [6]. Ontogenetic colour changes occur in association with development, independent of the environment, and are most commonly a response to different anti-predator requirements resulting from changes in life history [7]. The social significance of ontogenetic colour changes has received relatively little attention compared to the more dramatic short-term colour changes.
Ontogenetic colour changes are common in many crustaceans, typically in response to the changing camouflage requirements associated with changes in habitat or behaviour. Upon sexual maturation, female prawns (Leander serratus) develop white spots over their egg-bearing segments, possibly because the more vulnerable gravid females require better camouflage [8]. As isopods (Idotea montereyensis) grow they change from red to green to match the background as they move from rockpools filled with red algae to beds of green eelgrass [9]. Similarly, shore crabs (Carcinus maenus) exhibit white spots on their carapace when small, enabling them to blend into the mussel beds in which they live. As the crabs grow and move into algae beds, they lose their spots to better camouflage themselves [10]. In addition to the variation in colour pattern between juveniles and adults, individual crabs are also extremely variable [11].
Although responsible for some intraspecific variation in colouration, colour changes cannot explain the extreme phenotypic diversity in colour patterns found in many species. While the adaptive significance of short-or long-term colour changes is generally well known, the phenotypic diversity in colour pattern between individuals is often neglected. Whether the colour patterns are individually distinct or sexually dimorphic, as with ontogenetic colour changes, their adaptive significance is often interpreted in terms of predator avoidance [12][13][14][15]. However, there are a number of benefits associated with being individually recognisable [16,17]. To fully understand the development and function of colour pattern variation, it is necessary to first describe its morphological basis and the naturally occurring diversity [18].
Fiddler crabs (genus Uca) are often spectacularly coloured [19,20] and capable of dramatic colour changes. Their colour changes have been the focus of numerous studies, resulting in a description of rapid changes associated with courtship displays [21][22][23] and stress [19,24,25]. Some fiddler crabs also change colour as a thermoregulatory response to temperature changes [4,26] and to match the background [4,20,27]. Finally, at least 15 species have been found to exhibit rhythmic colour changes in association with their circadian rhythms, and the tidal and semilunar cycles [20,28,29]. Despite our detailed knowledge of the physiological processes behind these rhythmic colour changes, their adaptive significance remains a mystery. Furthermore, unlike many other crustaceans, fiddler crabs' ontogenetic colour changes have never been described. Uca mjoebergi do not appear to alter their colour and use the colour of their claws to recognise conspecific mates [30]. Species recognition requires a signal that is relatively uniform and stable across the population, whereas recognition of kin, neighbours or individuals requires distinct signals on an increasingly fine scale [31]. While numerous fiddler crab species exhibit extreme individual variations in colour and pattern [19,24,32], few studies have attempted to quantitatively describe this diversity [32] and none have examined their adaptive significance.
The endemic Australian fiddler crab Uca capricornis Crane, 1975 uses carapace colouration to discriminate neighbours from strangers [30]. Both sexes display a variety of colours on their black carapace, including: blue, yellow, orange and white. In order to function as cues of individual identity, these colour patterns should be relatively stable and include sufficient variation to reliably identify an individual. The aim of this paper is to quantitatively describe the carapace colour patterns of U. capricornis, including variation related to sex and size and short-and long-term colour changes.

Quantifying Colour Patterns
Females have a greater proportion of their carapace coloured than a similarly-sized male. The proportion decreases with a corresponding increase in the size of both males and females (GLM arcsine proportion carapace coloured: carapace width F 1, 272 = 76.6, p,0.0001; sex F 1, 272 = 50.9, p,0.0001; interaction F 1, 272 = 0.113, p = 0.7) (Figure 1 and 2). The result is that juveniles of both sexes have uniformly coloured carapaces, but the coloured area contracts in larger individuals, giving them a mottled pattern, before concentrating into 6 particular areas ( Figure 2). Crabs larger than approximately 12 mm generally exhibit some form of this pattern, consisting of a patch behind the eyestalks (the scarf), surrounded by another 5 spots. The exact size of these patches varies between individual crabs; from non-existent, to covering much of the carapace.
Juvenile crabs have the most variable colour patterns, due to their mottling (Figure 3b). Individuals larger than 12 mm vary in the size of the 6 specific areas (Figure 3b), with females' spots tending to be larger and more variable (8.3610.5 mm 2 , n = 82 ) than males' (2.763.3 mm 2 , n = 111) (Figure 3a).

Rapid Changes
When caught, or otherwise stressed, U. capricornis are capable of rapidly changing their colour. These short-term colour changes involved darkening their colour to some degree, based on human observation, and were more common in females (35 out of 43) than males (12 out of 22) (2-tailed Fisher's exact test, p = 0.04). Eighteen crabs darkened their entire colour pattern equally (Figure 4a). In another 17 crabs, the yellow scarf region darkened more noticeably than the white spots ( Figure 4b). While these crabs became steadily darker over the 20 minutes, the remaining 12 crabs darkened as normal but started to brighten, and in some cases almost regained their original colouration, within the 20 minutes (Figure 4c). In all cases the pattern remained unchanged, but did become more difficult to distinguish as the colours darkened.

Daily Changes
Uca capricornis do not regularly change their colouration over the course of a day. Of the 14 females and 13 males observed throughout a day, the vast majority (13 and 11 respectively) did not noticeably change their colour, and none altered their pattern.
The two males that did change brightened their colour slightly over the course of the day, peaking after low tide. In the larger male the change was most pronounced in the yellow scarf region (Figure 5a), while the smaller black and white male brightened all over ( Figure 5b). On the other hand, the single female that did change darkened after low tide (Figure 5c), apparently in response to being evicted from her burrow by another crab. These changes appeared quite subtle compared to the rapid changes described above.

Changes with the Semilunar Cycle
Uca capricornis do not rhythmically change their colour or pattern between moults in association with the semilunar cycle. No crabs noticeably changed their pattern or from one colour to another, and 12 out of 22 did not alter their colour in any way from one day to the next over a two week cycle. The remaining 11 crabs brightened or darkened their existing colours slightly from one day to the next. However, the number of changes made by each crab over the 14 day cycle varied, from 1 change (3 crabs), 2 changes (2 crabs), 4 changes (1 crab), 5 changes (3 crabs), 6 changes (1 crab), or 7 changes (1 crab). There was no apparent trend in the number of crabs either brightening or darkening with respect to the time in the cycle; there was a low level of both brightening and darkening, throughout the 2 weeks ( Figure 6).

Changes Associated with Moulting
On average, U. capricornis grew 1.6 mm with every moult (standard deviation = 0.5 mm, n = 37). The difference in pattern between individuals of different sizes is clearly the result of changes within individuals associated with moulting, as the correlation between colour pattern and size is consistent with the changes observed in moulting individuals. With each successive moult the pattern appears to contract and become more distinct, especially in males ( Figure 7).
In addition to the considerable changes in pattern, both sexes also changed the specific colour of their spots. The colour changes associated with moulting, as with changes in pattern, tend to follow a certain progression ( Figure 7). The first few moults of both sexes are a uniformly brilliant light blue, which pales to white over successive moults. They retain this black and white colouration over the next 3-5 moults, as they develop the mottled pattern and the beginnings of the distinctive scarf and spots. Upon reaching about 12 mm males and females start to differ.
In males the scarf and the two spots at the corners of their carapace turn yellow, while the lower spots generally remain white (Figure 7a). In addition to the yellow on their carapace, males over 12 mm also develop distinctive blue meral spots on their fourth pair of walking legs, which are retained by the largest individuals that have otherwise lost all carapace colour. Females larger than about 12 mm also develop more colourful carapaces, although they are more variable than those of males: their scarf and surrounding spots can be any combination of yellow, white, or blue ( Figure 7b). As with males, larger females can also develop blue meral spots however, these are not restricted to the last pair of legs, and may cover the entire leg.
After developing a yellow scarf, males do not change the colour of their patches again. The coloured areas steadily shrink over successive moults, until they disappear entirely (Figure 7a). On the other hand, although the colour and pattern of the 6 caged females large enough to develop colour remained relatively stable over successive moults, when they did change they were more variable and less likely to follow a certain progression (Figure 7b).

Perception of Colour Patterns
Based on the theoretical visual acuity of U. capricornis, the spacing between the receptive fields of ommatidia would be 0.9 mm for a viewing distance of 50 mm, 1.7 mm for 100 mm, and 2.6 mm for150 mm. The configuration of their typical colour patches appears to enable them to be detected by individual, nonadjacent ommatidia from as far away as 15 cm (Figure 8). This should enable U. capricornis to perceive the variation in colour pattern in both females ( Figure 8a) and males (Figure 8b).

Discussion
The colour patterns on the carapaces of Uca capricornis are extremely variable, although they do follow a certain progression with size and sex. Both males and females change from a uniform blue colour to a pattern of distinct white, yellow, or blue patches on a black background. The coloured patches of both sexes tend to shrink as the crabs grow, but females are generally more variable, with a wider variety of colours in larger patches than similarlysized males.
Habitat partitioning and differential predation pressure are linked to sexually dimorphic colour patterns in other crustaceans, such as the isopod Idotea baltica [13,14] and are involved in maintaining colour polymorphism in many species, including vertebrates [12]. There is no apparent habitat partitioning in U. capricornis based on sex, size, or colour morph. However, the different colour classes do appear to behave differently as larger individuals are more inclined to leave their burrows and wander through the colony (T Detto & PRY Backwell unpublished data). This behaviour may expose them to a greater risk of predation, which may select for their less colourful appearance. However, this cannot be proven until we identify their predators and determine their visual capabilities.
A likely explanation for the extreme variation in colour patterns in U. capricornis is their involvement in individual discrimination. Males live in close association with their neighbouring females (T Detto & PRY Backwell unpublished data) and use carapace colour patterns to distinguish between their female neighbour and unknown females [30]. The more variable colour patterns of females appear to be unique and may enable their mates to distinguish them from other females. Models of the theoretical perception of the average colour pattern suggest that the crabs are physiologically capable of discriminating variation in spot size and distribution. Recent work has also shown that another species, U. mjoebergi, possess colour vision [33], which may also mean U. capricornis can make use of the individual variation in colour. In addition to the ability to perceive variation in colour pattern, an individual's colour pattern needs to be relatively stable over time to function as an individual identifier, as colour changes may result in misidentification. A number of fiddler crabs reportedly change their colour around spring tide and over low tide, when waving and courtship activity is at a maximum [19,22,23,25]. We found no evidence that U. capricornis regularly change their colour in association with the semilunar or tidal cycle. The few crabs that did change their colour during the semilunar and tidal cycle were likely doing so in response to an unobserved social stimulus rather  than an endogenous rhythm. Uca capricornis are capable of rapidly changing their colour in response to capture or losing their burrow. This colour change involves darkening existing colours, probably by temporarily concentrating the coloured pigments and/or dispersing black pigment. The pattern remains unchanged and all evidence suggests that the crabs regain their original colouration. It is difficult to say what adaptive significance these rapid colour changes have.
They may be used in intraspecific communication, as a form of camouflage for crabs that have lost their burrow, or they may simply be a physiological response to stress.
While U. capricornis are clearly capable of rapid physiological colour changes, these do not appear to play a major role in the crabs' general colouration. Their colouration is determined by ontogenetic colour changes during moulting. Between moults, their colour pattern is very stable. They seem unable to change the position, number, or colour of their chromatophores between moults. While they are able to dull their colour when stressed, they then return to their original state. The carapace colour patterns of U. capricornis are thus stable enough to allow individual discrimination: they remain the same during an entire inter-moult period, which is likely longer than pairs remain together (T Detto & PRY Backwell unpublished data). Furthermore, larger females maintain a relatively similar colour pattern over several moults.
In addition to individual identity, the ontogenetic colour changes in U. capricornis potentially provide information about the sex, size, and reproductive status of the individual. The crabs do not develop their yellow colouration until growing to about 12 mm. This may be because they need time to accumulate the necessary carotenoids, or it may be a signal of sexual maturity. Female prawns and shrimps develop white chromatophores at sexual maturity [8]. The steadily decreasing size of the coloured patches in larger males also has the potential to signal competitive ability.
When first described, Uca capricornis was believed to be 2 distinct species: U. capricornis and U. pavo, based on the extreme difference in colouration between males of different sizes [34]. Together with the potential social significance of colour in U. capricornis, this highlights the importance of studying ontogenetic colour changes and our lack of understanding of their possible significance.

General Methods
This study was conducted on a large population of Uca capricornis in the vicinity of the mangrove boardwalk in the East Point Reserve, Darwin, Australia (12u249350 S, 130u509000 E). Several experiments were conducted on crabs housed within cages built on the mudflat, these cages are discussed in detail in the final section (changes associated with moulting).
Every effort was taken to minimise any possible effect of our presence on the behaviour of the crabs, including those in the cages, as they undergo a dramatic colour change when directly handled. Experience has proven Uca capricornis to be particularly resilient to human presence, and within 5 minutes of motionless observation the crabs resumed an apparently normal range of behaviour, often next to or even on top of the observer's feet. From this we have to assume that our presence had little impact on the behaviours reported below. Unfortunately, this also required us to rely on the human visual system to describe many of the long-term colour changes

Quantifying Colour Patterns
Colour pattern refers to the configuration of any coloured areas on the black carapace, regardless of their spectral composition. To quantify the carapace colour patterns of U. capricornis we caught and measured the carapace width of all individuals active on the surface within 18 plots (4 m 2 each) and a random sample of larger crabs, which were relatively sparse within the plots. We thus collected a total of 117 females and 159 males. We temporarily restrained them before taking a digital photograph of their carapace. Afterwards, the crabs were released unharmed.
All digital photographs were taken using manual white balance and included a colour standard. We used the manual colour correction function of Paint Shop Pro version 7.02 (Jasc Software 2000) to standardise the colour of the photographs and account for lighting differences. This allowed more accurate comparisons of individual differences and colour changes within individuals.
Using a raster-based graphics software [Paint Shop Pro version 7.02 (Jasc Software 2000)] we converted each crab carapace into a stylised black hexagon, with corners at the base of the last pair of legs, the widest points of the carapace, and the base of the eyestalks. We then converted any colour markings on the carapace to white before standardising the size and orientation of the images using ENVI software (Research Systems Colorado) to warp them to a standard carapace 45 mm wide and 25 mm high. The resulting images showed the relative position of the colour pattern (in white) on a black carapace.
We quantitatively examined the relationship between colouration, carapace size, and sex. Using the public domain Scion Image (Alpha 4.0.3.2) program (Scion Corporation), we measured the total size of the coloured area and the area of the carapace, using the measured carapace width as a scale. We then determined the proportion of the carapace that was coloured and examined its relationship with carapace size and sex using a generalised linear model. The data were tested for normality, and the proportion of the carapace that was coloured was arcsine transformed to conform to the assumptions of normality. Furthermore, we calculated the average size of the various colour patches by dividing the total coloured area by the number of distinct spots. Means are reported with the standard deviations.
To visualise the differences in colour pattern between the sexes and crabs of different sizes, we constructed images illustrating the probable colour patterns of various groups of crabs, including: males and females in general, and males and females within various size classes; ,8 mm, 8-9.9 mm, 10-11.9 mm; 12-13.9 mm; 14-15.9 mm; 16-17.9 mm; and males .18 mm (no photographed females were .18 mm). We overlayed a 161 mm grid on each of the warped images and assigned a value of 1 to each coordinate that was more than one third white and 0 to all other, uncoloured, coordinates. We then determined the proportion of the crabs that had colour at each coordinate and converted these percentages into RGB lightness values, with 0 (black) equivalent to no crabs coloured in that area and 255 (white) equivalent to all crabs coloured in that area. We used these values to colour the appropriate coordinates on a standardised carapace, producing an image that represents the likelihood that any particular area is coloured. These images are the equivalent of an average crab's colour pattern. To more clearly illustrate the variation in colour pattern between the groups, we constructed similar images using the standard deviation of crabs coloured at each coordinate.

Rapid Changes
We examined the ability of 65 randomly caught crabs (21 males and 44 females) to change colour over 20 minutes. We measured the spectral reflectance of various coloured areas on their carapace relative to a white 'Spectralon' standard with a USB2000 UV-VIS portable spectrophotometer (Ocean Optics Inc, Dunedin, FL, USA). The first measurements were taken as soon as possible after capture, and further measurements were taken of the same areas 5, 10 and 20 minutes after the initial measurements. After each set of measurements we also took a digital photograph. Between measurements, the crabs were kept in the shade in a clear plastic cup with 1 cm of seawater.

Daily Changes
We also examined whether U. capricornis change colour in relation to the tidal cycle. On the day after spring tide, over several semilunar cycles, we observed 27 crabs (13 males and 14 females), including those within the cages and several outside. We took digital photographs of each crab 2 hours to 1 hour before low tide (before), in the 1 hour around low tide (during), and 1 hour to 2 hours after low tide (after). Each crab thus had three photographs taken over a single day, each separated by 1.5 hours. We then examined the photographs be eye for any changes in colour or pattern based.

Changes with the Semilunar Cycle
During one semilunar cycle, from one neap tide to the next, we took digital photographs of all 22 crabs (10 males and 12 females) active in the cages at approximately low tide every day. We examined the photographs and identified the proportion of the crabs that were darker, brighter, or the same colour as the previous day to determine whether they underwent any rhythmic colour changes. As we only observed them over a single cycle we cannot make any definitive statements about their colour changes associated with the semilunar cycle, but can identify any obvious trends.

Changes Associated with Moulting
We constructed 5 circular cages (1 m in diameter) out of flyscreen mesh and buried them 10 cm in the mud within the U. capricornis population. The cages protruded a further 20 cm above the mud and a laminate strip glued around the inner and outer top 5 cm prevented crabs from climbing into or out of the cage. We removed all existing crabs from the cages and restocked each cage with 4 crabs; a small male and female and a large male and female. We measured the crabs, took a digital photograph of their carapace and superglued a coloured sequin to the posterior region of their carapace. Checking on the crabs every week, we identified and photographed those lacking a sequin, indicating they had recently moulted, before measuring and remarking them. Over the 4 months of the experiment we had to replace several of the crabs that disappeared, and although several reappeared, they were distinguishable by their size, colour pattern and the colour of their sequin. In total we observed and photographed 11 females and 10 males through at least one moult.

Perception of Colour Patterns
Finally we examined whether Uca capricornis are anatomically capable of perceiving the different colour patterns, by using a theoretical inter-ommatidial angle of 1u [35,36] as an estimate of the crabs' maximum spatial resolution. We calculated the spacing between the centres of adjacent ommatidia for distances of 50, 100, and 150 mm.
Centre spacing~Viewing distance|tan(0:5 0 ) We then assigned the standardized image of an average carapace (calculated while quantifying the colour pattern) a width of 14 mm, and projected this image on the crab's ommatidial array ( Figure 8).The crabs should be able to discriminate the various colour patches if they are perceived by individual, nonadjacent ommatidia.